Sandwich Hybridization Assay for HABs Detection and Monitoring: History
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Ping Gong
,
Anna K. Antrim
,
Sarah R. Bickman
,
Emily G. Cooley
,
Seung Ho Chung

As cyanobacterial harmful algal bloom (cHAB) events increase in scale, severity, frequency, and duration around the world, rapid and accurate monitoring and characterization tools have become critically essential for regulatory and management decision-making. The composition of cHAB-forming cyanobacteria community can change significantly over time and space and be altered by sample preservation and transportation, making in situ monitoring necessary to obtain real-time and localized information. Sandwich hybridization assay (SHA) utilizes capture oligonucleotide probes for sensitive detection of target-specific nucleic acid sequences. As an amplification-free molecular biology technology, SHA can be adapted for in-situ, real-time or near real-time detection and qualitatively or semi-quantitatively monitoring of cHAB-forming cyanobacteria, owing to its characteristics such as being rapid, portable, inexpensive, and amenable to automation, high sensitivity, specificity and robustness, and multiplexing (i.e., detecting multiple targets simultaneously). Despite its successful application in the monitoring of marine and freshwater phytoplankton, there is still room for improvement. The ability to identify a cHAB community rapidly would decrease delays in cyanotoxin analyses, reduce costs, and increase sample throughput, allowing for timely actions to improve environmental and human health and the understanding of short- and long-term bloom dynamics. Real-time detection and quantitation of HAB-forming cyanobacteria is essential for improving environmental and public health and reducing associated costs.

  • cyanobacteria
  • harmful algal bloom (HAB)
  • sandwich hybridization assay (SHA)
  • nucleic acids

1. Introduction

Harmful algal blooms (HABs) are characterized by increased phytoplankton biomass, declined dissolved oxygen, and sometimes by the production of cyanotoxins [1]. In freshwater, HABs tend to be dominated by cyanobacteria. Cyanobacterial harmful algal blooms (cHABs) are becoming more common around the world due to excess nutrient loads, eutrophication, and the changing climate [2]. cHABs, particularly Microcystis-dominated blooms, have been observed in every continent except Antarctica and have significant economic consequences. For example, 20 years ago, cyanobacterial blooms caused annual economic losses of up to $82 million on public health, fishery, and tourism in the U.S. alone [3]. cHABs are formed by a large number of genera, including but not limited to Microcystis, Anabaena/Dolichospermum, Aphanizomenon, Cylindrospermopsis, Planktothrix, Lyngbya/Microseira, and Phormidium/Microcoleus.
It has been extensively reported that cHAB-forming cyanobacteria can produce hundreds of metabolites known as cyanotoxins that are harmful to exposed aquatic and terrestrial animals, including humans [1][4]. For instance, Microcystis spp., Anabaena/Dolichospermum spp., and Planktothrix spp. are able to synthesize the peptide hepatotoxins, microcystins [5]. Cylindrospermopsis spp., Aphanizomenon spp., Anabaena spp., and Lyngbya wollei can produce hepatotoxic cylindrospermopsins [6]. Neurotoxic saxitoxins can be produced by Anabaena/Dolichospermum spp., Aphanizomenon spp., Cylindrospermopsis raciborskii, Planktothrix spp., and Lyngbya wollei [6][7]. Anatoxins, another class of neurotoxins, are produced by Aphanizomenon spp., Anabaena/Dolichospermum spp., and Phormidium spp. [8]. Notably, some of these genera, such as Anabaena/Dolichospermum spp., have the potential to produce all common classes of cyanotoxins. Although cyanotoxins-coding genes are often associated with HABs events, the detection of such genes does not explicitly imply the presence of cyanotoxins in the bloom [9].
A wide variety of approaches and technologies have been developed for the detection and monitoring of HABs-forming cyanobacteria, ranging from microscopic enumeration, analysis of Chlorophyll a, ATP and phycocyanin, quantitative polymerase chain reaction (qPCR), next-generation sequencing (NGS), enzyme-linked immunosorbent assays (ELISA), and high-pressure liquid chromatography (HPLC), to hyperspectral imaging, remote sensing, automated cell imaging systems, and machine learning [10]. When HABs occur, there are many questions, including whether a HAB event has occurred, what cyanobacteria are present and caused the event, whether and what cyanotoxins are released, and how dynamic the cyanobacteria and cyanotoxins exist. Many tools are necessary to answer these questions, and there is currently a lack of portable and field-deployable tools for in situ real-time identification and monitoring of cyanobacteria in the waterbody of concern. In situ, real-time monitoring is preferred, since there is a loss of sample representation during sample collection, transportation, and storage [11]; and cyanobacterial community composition can change spatially and temporally. Therefore, there is a need for rapid, real-time, and reliable tests that can provide local or in situ answers for cyanobacterial monitoring.

2. What Is a SHA?

Hybridization is a process of two complementary single-stranded DNA or RNA molecules forming a single double-stranded molecule through base pairing. Sandwich hybridization assay (SHA) is a molecular technique based on successive hybridization of two oligonucleotide probes: a capture probe used to immobilize the target DNA or RNA on a solid support and a signal probe labeled with a detectable marker to quantify the target copy number [12]. A segment of the target molecule is “sandwiched” between the two probes in this amplification-free, direct capture method [13]. In general, the capture probe is designed with a high specificity to the target cyanobacteria, whereas the signal probe hybridizes to a conserved sequence found in target gene and is often labeled with either a fluorophore or a digoxygenin (DIG) to obtain a measurable signal. Considerations in capture probe design include but are not limited to the following: specificity to target organisms, proximity to signal probe (within 100–250 bp), <70% similarity with signal probe, GC content (i.e., percentage of guanine and cytosine in a DNA or RNA molecule) between 40% and 60%, melting temperature between 69 °C and 74 °C, secondary structure stability less than 34 °C, and homodimer stability less than 17 °C [14]. When a signal probe is labeled with a fluorophore (e.g., Cy5), a fluorescence reader is used to read out the target molecules trapped by the capture probe [15][16]. For DIG-labeled signal probes, anti-DIG antibodies in conjugation with horseradish peroxide (HRP) or alkaline phosphatase (AP) are added along with such a substrate as 3,3′,5,5′-tetra-methylbenzidineto (TMB) or 2′-(2-benzothiazolyl)-6′-hydroxybenzothiazole phosphate (BBTP) to produce a colorimetric or fluorescent readout [12][14][17].

3. SHA Development and Application

The SHA was initially developed in the late 1970s for mapping transcripts to the genome of adenovirus type 2 [18]. It was nearly twenty years later when the first U.S. patent for this technology was awarded in 1995 to Jennifer K. Ishii and Soumitra Ghosh, both with Siska Diagnostics Inc., La Jolla, CA. The two inventors developed a two-step sandwich hybridization technique for detection of as little as 10−17 moles of nucleic acid molecules in solution, without requiring the use of radioactive compounds: the target nucleic acids are captured by hybridization with oligonucleotides covalently attached to a polystyrene solid support to form complexes that are then hybridized to detection oligonucleotides [19]. Three years later, Tyagi et al. [20] were granted another U.S. patent for multiple SHA background “noise” reduction measures, such as the use of separate capture and signal/reporter probes, separation from immobilized capture probes by cleavage and isolation, use of RNA binary probes and an RNA-directed RNA ligase, and amplification by an RNA directed RNA polymerase.
The SHA protocol has been modified to meet specific requirements for a wide range of application. For instance, the solid support has evolved from membrane materials in early times [21] to nylon beads [22][23], magnetic beads [12][17][24][25], polystyrene prongs [14][19], microarray glass slide [16], and microtiter plates [24][26]. Due to their physical porous structure, membranes give high background and steric constraints, causing difficulties in quantitative analysis [21][24]. Beads possess a large surface area with a well-defined capacity and rapid binding kinetics, leading to a higher fixation capacity and a higher hybridization yield, whereas microtiter plates and microarrays are better adapted for the simultaneous handling of a large number of samples [16][24][26]. In order to immobilize a capture probe to a solid support, the support surface may be coated with streptavidin to which the biotinylated capture probe is bound [12][14][17][22][23][25]. Alternatively, the capture probe can covalently bind to aminated solid surface via a carbodiimide crosslinker [16][24][26]. The use of unlabeled “helper” probes (between capture and signal probes) increased hybridization efficiency by 15- to 40-fold [17][27]. Instead of Cy5-, Cy3-, or DIG-labelling at one end, double DIG-labeling at both the 5′- and 3′-ends of detection probes significantly enhanced signal intensity [14].
The synthetic DNA mimic peptide nucleic acid (PNA) have been used to replace DNA probes [15][26] because of its superior hybridization characteristics and improved biochemical properties, including resistance to enzymatic degradation, increased sequence specificity to complementary DNA, and higher stability when bound with complementary DNA [26][28]. Although the reported limit of detection (LOD) for miRNA using PNA probes was 10 nM corresponding to 2 × 1011 target molecules in a 30 μL sample vial (at least 200-fold higher than using DNA probes), million-fold increases in target concentration can be realized to achieve the theoretically ideal LOD for low abundance miRNA targets on the order of 104 targets, 7 orders of magnitude lower than the reported LOD [15]. For non-miRNA targets, a cyclopentane-modified capture PNA probe (PNAα) in combination with a biotin-labeled signal PNA probe (PNAβ), a commercially available polymer of Horse Radish Peroxidase-avidin (poly-HRP-avidin) and tetramethylbenzidine (TMB), can detect 10 zmol (1 zeptomole = 10−21 mole or 10−6 femtomole (fmole)) of target DNA [26], which is 3000-fold lower than the 0.03 fmole of LOD [16][24], the lowest.
From the instrumentation perspective, a regular SHA would not require anything but visual inspection if a qualitative endpoint of color change is measured or would simply use a plate reader that measures quantitatively either fluorescence (e.g., at excitation wavelength 430 nm and emission wavelength 560 nm) [12][17] or absorbance (e.g., optical density at 450 nm) [14][24][26]. Other sophisticated equipment, such as liquid scintillation counter and SERRS, have been employed when the detection probe is labeled with [35S]-ddATP [24] and rhodamine 6G dye [25], respectively. A capillary electrophoresis running nonionic surfactant micelle-containing buffers was used to separate the sandwich complex (i.e., a target DNA sandwich-hybridized with a γ-substituted PNA amphiphile (γPNAA) probe and a DNA probe) from unbound γPNAA probes, DNA probes and target DNAs via a mobility shift assay [15]. A microarray scanner is needed when capture probes are printed on a glass slide [16].
Typical SHA protocols can be completed within 3–4 h and often use saline-sodium citrate (SSC, e.g., 1× SSC made of 0.15 M NaCl and 0.015 M sodium citrate) as the main component of hybridization and washing buffers. Compared with SSC/NaCl-based buffers, GuSCN (guanidine thiocyanate) is another commonly used base reagent in hybridization buffers [14][22][23] because it is effective at disrupting cells, inactivates nucleases, and permits direct, specific hybridization at much lower temperatures [29][30]. Other widely used ingredients in these buffers include Tween 20, polyvinyl pyrrolidone (PVP), ethylenediaminetetraacetic acid (EDTA), sodium dodecyl sulfate (SDS), Tris, formamide, maleic acid, and dextran sulfate.

4. SHA Application to Cyanobacterial Detection and Monitoring

Although SHA has been around for nearly five decades, its application to cHABs detection and monitoring, began only two decades ago, which was likely driven by the rising demand for molecular technologies enabling in situ identifying, sensing, and monitoring HAB-causing cyanobacteria [31]. Surveies identified seven peer-researchs and one thesis publication. Matsunaga et al. [32][33] designed the first set of capture probes targeting the genus specific region of the 16S rRNA sequences from five cyanobacterial genera (Anabaena, Microcystis, Nostoc, Oscillatoria, and Synechococcus). These probes were immobilized on bacterial magnetic particles (BMPs) isolated from the magnetic bacterium Magnetospirillum magneticum AMB-1 via streptavidin-biotin conjugation. A DIG-labeled cyanobacterial universal probe CYA781R was used as the signal probe [34]. An anti-DIG-AP antibody was used for signal detection after addition of the CDP-Star™ Substrate with Emerald-II™ Enhancer or the AttoPhos® AP substrate. Results demonstrated high discriminatory power of the genus-specific capture probes, which produced significantly higher fluorescence when hybridized to the 16S rRNA amplicons from the strains belonging to their respective target genus [32]. The researchers further automated the entire hybridization and detection process using a magnetic separation robot and transformed the assay into a 96-microwell format to increase the throughput [33].
Castiglioni et al. [35] developed a microarray spotted with universal oligo probes (i.e., 5′ NH2-modified “zip code” oligonucleotides carrying a poly (dA)10 tail at their 5′ ends covalently immobilized on a CodeLink slide) to profile the abundance and diversity of 19 cyanobacterial groups identified by phylogenetic analysis of 338 sequences of cyanobacterial 16S rRNA genes. Prior to array hybridization, a 30-cycle sandwich hybridization (called ligation detection reaction, LDR) was performed in a thermal cycler with an LDR mixture made of a discriminating probe labeled with Cy3 dye at the 5′ end, a common probe phosphorylated at the 5′ end and carrying a czip code (i.e., oligos complementary to the “zip code” probe) at the 3′ end, a DNA ligase, and a purified PCR product of cyanobacterial 16S rRNA. Each pair of discriminating probe and common probe (excluding the czip code) was designed specifically to target one of the 19 cyanobacterial groups. Array hybridization was carried out in a dark chamber at 65 °C for 1 h. After post-hybridization washing and drying, the Cy3 green fluorescence intensity was acquired for array spots using a laser scanner with settings of λex = 543 nm and λem = 570 nm. This SHA-based microarray approach was validated by testing 95 known 16S rRNA amplicons of single strain (24 from axenic strains, 27 from isolated strains, and 44 from cloned fragments recovered from lake samples), unbalanced mixtures of different known 16S rRNA amplicons, and an unknown environmental sample, all of which demonstrated a high discriminative power and sensitivity (LOD = 1 fmole).
The above-mentioned three early studies only evaluated PCR products of 16S rRNA in pure cultured or environmental cyanobacteria. Later studies directly analyzed non-amplified nucleic acids extracted from pure cyanobacterial cultures or environmental samples. Zhu et al. [36][37] designed two pairs of Microcystis-specific capture and signal probes, one targeting the PC-IGS (phycocyanin intergenic spacer) region [38] and the other targeting the mcyJ gene. These probes not only qualitatively discriminated Microcystis from other cyanobacterial genera (Anabaena, Aphanizomen, and Planktothrix (Oscillatoria)) but also quantitatively detected Microcystis populations at environmentally relevant densities as low as 100 cells/mL. These SHA results were validated by microscopic enumeration technique [36][37]. Following the method of Goffredi et al. [14], Dearth and coworkers [39][40] also designed a Microcystis-specific capture probe (MIC593) and adopted the bacterial universal probe EUB338 [41] as the signal probe, both targeting the 16S rRNA gene. They modified the Goffredi method [14] by replacing the biotin-coated polystyrene prongs with a streptavidin-coated microwell plate, and the streptavidin-biotinylated capture probe with biotinylated capture probe. The modified SHA had a LOD of 1.5 × 104 cells/250 μL homogenate and a linear range between 7.75 × 104 and 1.30 × 106 cells/250 μL homogenate, corresponding to the cell density of a moderate Microcystis bloom (1.00 × 105 cells/mL). Using the modified method, the researchers investigated the influence of environmental factors (light intensity and temperature) on Microcystis populations.
Another major development in this field is the integration of SHA to an automated sampler such as Environmental Sample Processor (ESP). Motivated by the so-called “ecogenomic sensors” notion of using an ordered array of different probes to detect a variety of organisms in a single sample, a group of researchers in the Monterey Bay Aquarium Research Institute (MBARI) developed a novel SHA-based probe array as one of the analytical modules in the ESP [42]. A series of publications by this group documented the conceptualization, development, field demonstration, deployment, and refinement of ESP to meet the growing needs of in situ, real-time HABs investigation [30][31][43][44][45][46]. One of these publications [30] reported the use of target-specific SHA probes for successful detection of 16S rRNA indicative of marine cyanobacteria (Synechococcus) and other phylogenetically distinct clades of marine bacterioplankton in a 96-well plate format as well as low-density ESP arrays printed on a membrane support. Samples subjected to investigation included target and non-target products derived from in vitro transcription of 16S rRNA genes as well as extracted RNA from collected natural seawater. Reported detection limits were between 0.10–1.98 and 4.43–12.54 fmole/mL homogenate for the 96-well plate and array SHA, respectively.
More recently, Microbia Environment Inc. developed and patented CARLA (Cellular Activity RNA-based eLisA), a commercial biosensor technology based on the SHA with a detection probe coupled with an enzymatic activity that induces a colorimetric signal proportional to the quantity of sampled rRNA (https://www.microbia-environnement.com/en/technology/) (accessed on 20 July 2022). Species-specific detection and quantification of target cyanobacteria, such as Microcystis, Planktothrix, and the ADA clade (Anabaena/Dolichospermum/Aphanizomenon), can be accomplished in less than 3 h after RNA extraction from water samples.

5. Advantages of SHA

When used as a molecular tool for routine detection and monitoring of HAB-causing cyanobacteria, SHA has many advantages over conventional techniques such as light microscopy and quantitative polymerase chain reaction (qPCR). Traditionally, samples were collected and transported to a laboratory where microscopy was performed to identify and enumerate cyanobacterial communities. The performer requires special training and hands-on experiences in morphological observation and taxonomic identification or classification. This process is time-consuming (taking days or longer) and labor-intensive, and it may introduce personal bias. Even though such automated and field deployable cell imaging systems as Imaging Flow CytoBot (IFCB, https://mclanelabs.com/imaging-flowcytobot/) (accessed on 20 July 2022) and FlowCAM (https://www.fluidimaging.com/) (accessed on 20 July 2022) can quickly, accurately, and reliably identify and quantify cyanobacteria, they are currently cost-prohibitive, limiting their wider application for in situ detection and monitoring of cHAB species. In contrast, SHA is convenient to perform and appears to be rapid (a few hours), accurate (no or low cross-reaction with non-target species), repeatable, reliable, and highly amenable to automation (using robotic workstation) and multiplexing (in 96-well or array format) [30][33][35][39][40].
Although qPCR is more sensitive with a much lower detection limit (a few copies of target molecules) [47] and a much broader linear dynamic range (>5 orders of magnitude) [48], SHA does not require amplification and sophisticated equipment, and its readout may be visualized by color change or quantified using smaller, more portable, and much less expensive instrumentation, making it more cost-effective and field-deployable to track target species in near real-time and in situ [30][43][44][45][46]. Although new omics technologies (i.e., metagenomics, transcriptomics, proteomics, and metabolomics) are fast-growing and powerful tools with great potential applications for cyanobacterial community diversity and dynamics studies [49], such applications are currently limited to laboratory benchtop research [31]. In a comparative study for quantifying laboratory cultures of the ichthyotoxic raphidophyte Heterosigma akashiwo, Doll et al. [50]. observed a high degree of correlation between qPCR and SHA responses. With an LOD at 1 fmole of target molecules [35] or 100 cells/mL [36][37], SHA possesses a sensitivity required for detecting typical cyanobacterial populations observed in the early phase of blooms.
Other benefits of using SHA over other molecular methods include relatively low per sample cost, processing time, and capital investment on instrumentation. When integrated into an ESP, the cost of SHA is estimated to be between $7 and $10 per sample [42]. The hands-on time spent performing SHA is estimated to be 15–20 min and approximately 30–40 samples can be analyzed in an eight-hour day, when using an automated processor.

6. Technical Limitations of SHA

SHA is often considered a semi-quantitative assay [42] with a narrow linear range of two orders of magnitude, e.g., 5 × 102 to 2.5 × 104 cells/mL [36] or 1–100 pM of target nucleic acids [16]. Although the capture probes are capable of distinguishing between target sequences with as little a difference as a single base pair [15], such discriminatory power is dependent on the availability of sequence information on taxa of interest. This is mostly a concern in developing capture probes for a genus or a phylogenetic clade, which often target such ribosomal subunits as 16S, 18S, 23S, and 28S rRNAs. These rRNAs are highly conserved such that capture probes targeting these sequences risk cross-species hybridization and cannot separate closely related species of interest. A solution to this issue is to select unique or divergent genes in the target organism (e.g., mcyJ gene from Microcystis strains [36][37]). Moreover, SHA measures the copy numbers of target genes (e.g., 16S rRNA), which cannot be directly converted to cell density of target cyanobacterial strains without constructing a standard curve between cell counts and signal intensity or gene copy number. This is due to the fact that the expression level of assayed gene (copy number per gene) is affected by the cellular physiological status [40][50]. SHA results can be adjusted accordingly, when it is known how many copies of 16S rRNA are present in a cyanobacterial cell (e.g., Microcystis aeruginosa NIES-843 [51]).
Similar to other amplification- or hybridization-based techniques (e.g., regular PCR, qPCR, reverse transcription PCR, southern blotting, and northern blotting), cell lysis is an important preparatory step in SHA to release target intracellular molecules. The sensitivity and quantification accuracy of SHA are affected by the quality and yield of extracted nucleic acids. For cyanobacteria, some species are easier to lyse than others. For instance, Lyngbya spp. are typically difficult to lyse due to thicker cell walls and protective sheaths [52]. Many physical, chemical, and enzymatic methods have been employed to lyse cyanobacterial cells, however, there is no one method capable of disrupting all cyanobacterial species in a satisfactory fashion [53]. For instance, it was reported that xanthogenate, a polysaccharide solubilizing compound, was able to lyse a wide variety of cyanobacterial genera but with varying RNA yields [53]. Furthermore, xanthogenate and other common chemicals used for lysis may interfere with SHA reagents, leading to inaccurate results [54]. Although chemical lysis could cause interference with the SHA chemistry, repeated freeze-thaw cycles or mechanical bead beating in combination with a lysozyme can lead to more efficient RNA extraction [39][40][52][55].

This entry is adapted from the peer-reviewed paper 10.3390/bios12080640

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